bioRxiv preprint doi: https://doi.org/10.1101/193474; this version posted September 25, 2017. The copyright holder for this preprint (which was not certified by peer review) is the author/funder. All rights reserved. No reuse allowed without permission.
Architecture of mammalian centriole distal appendages accommodates distinct
blade and matrix functional elements
T. Tony Yang1, Weng Man Chong1, Won-Jing Wang2, Gregory Mazo3, Barbara Tanos4,
Zhengmin Chen1, Thi Minh Nguyet Tran1, Yi-De Chen1, Rueyhung Roc Weng1, Chia-
En Huang1, Wann-Neng Jane5, Meng-Fu Bryan Tsou*3, Jung-Chi Liao*1,6
1Institute of Atomic and Molecular Sciences, Academia Sinica, Taipei, 10617, Taiwan
2Institute of Biochemistry and Molecular Biology, National Yang Ming University,
Taipei, 11221, Taiwan
3Cell Biology Program, Memorial Sloan-Kettering Cancer Center, New York, NY
10065, USA
4Institute of Cancer Research, London, United Kingdom
5Institute of Plant and Microbial Biology, Academia Sinica, Taipei 11529, Taiwan
6Genome and Systems Biology Degree Program, National Taiwan University, Taipei,
10617, Taiwan
*Corresponding authors:
Jung-Chi Liao
Meng-Fu Bryan Tsou
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Abstract
Distal appendages (DAPs) are nanoscale, pinwheel-like structures protruding from the
distal end of the centriole that mediate membrane docking during ciliogenesis,
marking the cilia base around the ciliary gate. Here, we determined a superresolved
multiplex of 16 centriole-distal-end components. Surprisingly, rather than pinwheels,
intact DAPs exhibit a cone-shaped architecture with components filling the space
between each pinwheel blade, a new structural element we termed the distal
appendage matrix (DAM). Specifically, CEP83, CEP89, SCLT1, and CEP164 form
the backbone of pinwheel blades, with CEP83 confined at the root and CEP164
extending to the tip near the membrane-docking site. By contrast, FBF1 marks the
distal end of the DAM near the ciliary membrane. Strikingly, unlike CEP164 which is
essential for ciliogenesis, FBF1 is required for ciliary gating of transmembrane
proteins, revealing DAPs as an essential component of the ciliary gate. Our findings
redefine both the structure and function of DAPs.
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Introduction
All cilia grow from the distal end of centrioles/basal bodies 1-4. To initiate primary
ciliogenesis, a mature vertebrate centriole docks to the membrane, recruits membrane
vesicles, regulates axoneme growth, and forms part of the ciliary base that gates the
ciliary compartment, all of which depend on its distal end 5-9. In particular, distal
appendages (DAPs) are previously defined as nine-bladed, pinwheel-like structures
protruding from the distal periphery of vertebrate centrioles 10 and are thought to be
involved in many of the aforementioned steps of ciliogenesis. CEP83, CEP89,
SCLT1, CEP164, and FBF1 are core DAP components recruited to centriole distal
ends prior to and independently of ciliogenesis 5. These components are assembled at
the G2/M border, and assembly is dependent on C2CD3, another centriole-distal-end
protein 11. In response to cell cycle cues in G1 that stimulate ciliogenesis, other
proteins including TTBK2, recycling endosome components (Rab8, Rab11, and
EHD1), IFT88, and ARL13B are recruited to DAPs along with the associated
membrane vesicles 12-17. Upon membrane docking mediated by DAPs, centriole distal
ends are further modified to release negative regulators of ciliogenesis, such as CP110
or CEP97 18, which enables nucleation of the axoneme from the centriole. At the
nascent axoneme bud, a specialized structure known as the transition zone (TZ) is
built as part of the ciliary barrier from dozens of protein components recruited from
the cytoplasm 19, guided in part by the residential centriole-distal-end proteins
CEP290 and CEP162 20, 21. The continuous growth and maintenance of the axoneme
in the gated ciliary compartment are further supported by the intraflagellar transport
(IFT) machinery 22, generating a functionally intact primary cilium that serves as a
sensory organelle.
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By initiating ciliogenesis, DAPs are situated in a highly intricate multi-junction
position that marks the border between the centriole and axoneme (or TZ), as well as
the junction between the plasma membrane and the ciliary membrane 5-9. This unique
location raises the possibility that DAPs and their associated factors may also form
part of the structural barrier gating the ciliary compartment. However, the pinwheel-
like morphology, as defined by electron microscopy (EM), does not intuitively
suggest how DAPs can serve as a barrier, given that the large space between each
pinwheel blade would allow many substances to pass through with little or no
resistance. Thus, DAPs may simply function as the anchor mediating membrane
docking or IFT machinery recruitment. Alternatively, however, the pinwheel shape
may not morphologically reflect the full complexity of DAPs, due the limitations of
EM.
Here we resolved 12 DAP-associated molecular species and four additional proteins
surrounding DAPs using direct stochastic optical reconstruction microscopy
(dSTORM) 23, 24, revealing a 3D nanoscopic resolution of the protein megacomplex in
intact cells and reaching a level of detail not previously achieved by current structure
biology techniques. We found the architecture supports a previously unrecognized
structure that gates transmembrane proteins. Together, our findings reveal an
unprecedented architectural and functional framework at the base of mammalian
primary cilia.
Results
dSTORM superresolution imaging reveals differential radial occupancies of DAP
proteins
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To systematically characterize DAPs and centriole distal ends, our dSTORM imaging
analysis focused on core DAP components (CEP83, CEP89, SCLT1, CEP164, and
FBF1), DAP-associated proteins (Cby1, TTBK2, and IFT88), other centriole-distal-
end proteins (C2CD3, CP110, CEP97, CEP162, and CEP290), ciliary membrane
proteins (ARL13B and EHD1), and the subdistal appendage (sDAP) component
ODF2. Superresolution microscopy has been used previously to localize centriolar
proteins 25-29, but systematic determination of proteins at the centriole distal end has
never been performed. Here, mature centrioles oriented axially or laterally
(perpendicular or parallel to the imaging plane) in human retinal pigment epithelial
(RPE-1) cells were imaged. As depicted in Fig. 1a, visualization of axially and
laterally oriented centrioles, respectively, allows clear determination of the radial and
longitudinal distributions of centriolar components, leading to a 3D architectural map
of the centriole distal end.
Analysis of axially oriented centrioles revealed radial localization of core DAP
components exhibiting nine-fold symmetric ring-like patterns that differ in
size/diameter, with CEP164 and FBF1 forming the largest ring, followed by SCLT1,
CEP89, and CEP83 (Fig. 1b; Supplementary Table 1). CEP164 displays a broader
radial distribution with distinct outer and an inner intensity peaks (Fig. 1b and
Supplementary Fig. 1), resembling the pinwheel-like pattern characteristic of DAPs as
defined by EM (Fig. 1c). By contrast, C2CD3, a protein essential for DAP assembly,
occupies a surprisingly small, more compact region inside the centriole lumen than
CP110, CEP97, CEP290, and CEP162 (Fig. 1b, c), all of which form a ring-like
distribution of ~170200 nm in diameter (Fig. 1d, Supplementary Fig. 2;
Supplementary Table 1), similar in size to the centriolar axoneme (Fig. 1c). This
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suggests that C2CD3 is not part of the DAP slanted structure defined by EM; rather, it
is a luminal protein likely defining the distal environment of centrioles required for
appendage formation.
CEP83, CEP89, SCLT1, and CEP164 form a blade-like arrangement to which
FBF1 is not localized
Two-color dSTORM imaging of the radial distribution further revealed distinct
angular positions among the core DAP proteins (Fig. 2ae). Images of pairs of
CEP83-SCLT1 (Fig. 2a), CEP89-CEP164 (Fig. 2b), and CEP164-SCLT1 (Fig. 2c)
showed that they can all be allocated to tilted lines resembling the arrangement of the
nine electron-dense DAP blades observed in EM images. SCLT1 was often localized
within the blades of CEP164 propeller-like signals (Fig 2c; Supplementary Fig. 3). By
contrast, FBF1 fills the circumferential gaps of CEP164 puncta (Fig. 2d) as well as
SCLT1 puncta (Fig. 2e), likely localized to an open area between adjacent electron-
dense DAPs observed by EM. Histograms of angular spacing between two puncta,
one from each of a pair of DAP proteins collected from multiple cilia, revealed that
CEP83-SCLT1, CEP89-CEP164, and CEP164-SCLT1 pairs yielded peaks close to
multiples of 40, or 360/9 (Fig. 2f; Supplementary Fig. 4; Supplementary Table 3),
reflecting their similar circumferential arrangements. By contrast, histograms for
CEP164-FBF1 and FBF1-SCLT1 pairs revealed peaks close to 20, 60, etc (Fig. 2f;
Supplementary Table 3), confirming their alternating arrangements. Thus, unlike other
core DAP components, which localize to the slanted DAP blades (Fig. 2ac, g), FBF1
is instead associated with the space between DAP blades (Fig. 2de, g).
Combination of axial and lateral superresolution images reveals a 3D molecular
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map of DAPs
The lateral view, revealed by a series of two-color dSTORM images (Fig. 3a), showed
that CEP83, SCLT1, and FBF1 respectively form distinct thin layers along the
proximal-to-distal axis of the centriole (Fig. 3b). Consistent with its wide radial
distribution, CEP164 also occupies a broad longitudinal localization, forming a
trapezoid-like shape with two tapered sides, resembling that of the DAP slanted
structure observed by EM (Fig. 3b, d). This suggests that CEP164 forms the primary
backbone of DAPs. Meanwhile, CEP89 occupies two layers at the centriole distal end
both with a similar width but separated axially by ~130 nm (Fig. 3b; Supplementary
Fig. 5). The distal layer of CEP89 is close to the same longitudinal position as FBF1
(Fig. 3b; Supplementary Fig. 6), whereas the proximal layer is located longitudinally
close to ODF2 in sDAPs (Fig. 3c; Supplementary Fig. 6), suggesting possible dual
roles for CEP89 in DAPs and sDAPs. By overlaying the dSTORM signals over EM
images of ciliated centrioles, we found that CEP164, CEP83, SCLT1, and FBF1 align
well with DAPs, with FBF1 localizing closest to the ciliary membrane (Fig. 3d).
CP110 and CEP97 in non-ciliated centrioles are localized at a longitudinal position,
similar to SCLT1 (Fig. 3e). Collectively, the relative positions of DAP proteins in the
lateral view reflect the contour of a DAP blade (Fig. 3f).
Combining information from all relative axial and lateral positions, we built a 3D
computational model to visualize the positioning of proteins at the DAP region (Fig.
3g), with a backbone structure illustrated based on EM 10. CEP164, CEP83, SCLT1,
and the distal layer of CEP89 mapped well onto the 3D-reconstructed DAPs (Fig. 3h).
By contrast, FBF1 localizes to the distal regions of the DAP gaps between adjacent
CEP164 molecules, possibly near or on the proximal boundary of the ciliary pocket
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(Fig. 3i). Thus, FBF1 may serve a role distinct from other core DAP components.
ARL13B lies adjacent to DAPs during ciliary initiation but is excluded from the
proximal transition zone upon ciliary maturation
Building upon this 3D nanoscale map of DAPs, we continued to add DAP-associated
proteins to the map. We first examined the spatial relationship between DAPs and the
membrane. ARL13B is a small GTPase associated with the ciliary membrane and
ciliary vesicles 30, 31. During the early stages of cilia growth (following an 8 h serum
starvation), ARL13B puncta were localized in close proximity to FBF1 puncta (Fig
4a, b). However, after cilia had grown fully (after a 24 h starvation), ARL13B signals
occupied the entire primary cilium, similar to IFT88 signals (Supplementary Fig. 7),
but were completely excluded from the ciliary base above DAPs where the proximal
TZ was located (Fig. 4c; Supplementary Fig. 8). These results suggest that ciliary
vesicles carrying ARL13B may initially dock at the DAP region, but are later
excluded from at least the proximal end of the TZ of intact cilia, consistent with the
previous observation of a ciliary zone of exclusion (CIZE) 32. EHD1, a protein
involved in ciliary vesicle formation, was found to occupy a space wider than the
diameter of the primary cilium (Fig. 4d; Supplementary Fig. 8 and 9; Supplementary
Video 2), consistent with its presence on the ciliary pocket membrane 15.
We also examined the localization of TTBK2, a kinase interacting with CEP164 and
promoting ciliogenesis 12, 33. In proliferating cells, TTBK2 was sporadically scattered
with an asymmetric and incomplete coverage around the centriole (Fig. 4e). However,
upon serum starvation, a broadened radial and longitudinal distribution of TTBK2
was induced, forming a thicker occupancy covering the distal regions of DAPs close
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to SCLT1 (Fig. 4eg; Supplementary Fig. 10).
IFT88 and FBF1 localize to the distal appendage matrix, a newly identified
region between the nine distal appendage blades
We next examined IFT88, a core IFT complex component known to be associated
with DAPs 25. We found that IFT88 occupied a broad angular distribution without
clearly distinguishable nine puncta in the axial view (Fig. 5a). Lateral views revealed
that in addition to being distributed along the entire ciliary axoneme, IFT88 is heavily
concentrated at the ciliary base, at a longitudinal position spanning and proximal to
the SCLT1 molecules (Fig. 5b, c, and Supplementary Fig. 11). To further uncover the
exact localization of IFT88 close to DAPs, we performed 3D two-color dSTORM
imaging of IFT88 and SCLT1. We found that IFT88 occupied a conical distribution
(Fig. 5d, e; Supplementary Fig. 12; Supplementary Video 1), resembling the tapered
shape of DAPs. Strikingly, the relative distributions of IFT88 and SCLT1 reveal
abundant IFT88 proteins filling the gaps between neighboring SCLT1 molecules (Fig.
5f), demonstrating that molecules can occupy or be retained in this unexplored area
between DAP blades. We therefore designated the regions between the nine DAP
blades as the distal appendage matrix (DAM), in which proteins such as IFT can be
concentrated. In this sense, the intact DAP structure should be viewed as a conical
addition at the centriole distal end consisting of the distal appendage blades (DABs)
and matrix (DAM) (Fig. 5g).
The positioning of FBF1 at DAP gaps toward the ciliary membrane suggests that
FBF1 localizes at the distal end of the DAM interface with the ciliary membrane (Fig.
5h). Indeed, two-color IFT88-FBF1 dSTORM images showed that IFT88 coexists at
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the angular positions of FBF1 (Fig. 5i), and histograms of angular spacing revealed
peaks close to multiples of 40° (Fig. 5j), confirming the positioning of FBF1 at the
DAM. To check whether FBF1 and IFT88 at the DAM are functionally related, we
generated the FBF1-/- CRISPR knockout line (Supplementary Fig. 13) and examined
IFT88 localization in this region. Notably, in FBF1-/- cells, the distribution of IFT88
was reduced to a size comparable with that of CEP83 (Fig 5k, 5l), suggesting that
IFT88 cannot span the DAM region without FBF1. The causal effect of FBF1
depletion on IFT88 distribution is consistent with the observation of interactions
between FBF1 and IFT88 in a previous study 34. Our results potentially imply a
functional role for FBF1 in maintaining the DAM spatial arrangement (Fig. 5m).
We next tested whether DAB components perform a role in the structural arrangement
of the DAM, and vice versa. Axial-view dSTORM imaging revealed no obvious
change in CEP164 distribution upon FBF1 knockout (Fig. 6a, c). By contrast,
knockdown of CEP164 did reduce the radial distribution of FBF1 (Fig. 6b, d). Thus,
without CEP164 at the DAB tips, even though FBF1 was recruited to the DAP region
5, it was unable to reach the outermost DAM location (Fig. 6e). CEP164 depletion has
been shown to affect IFT recruitment 35, and it is also possible that the DAM was
damaged upon knockdown of CEP164, resulting in the collapse of the FBF1 ring.
Subsequently, we expanded our 3D computational model to include all ciliary
structures examined, including the membrane, pocket, axoneme, DABs, and the DAM
(Fig. 6f, g; Supplementary Fig. 14; Supplementary Video 3).
DAB proteins but not the DAM protein FBF1 are essential for ciliogenesis
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We then examined whether DABs and the DAM have distinct functions in
ciliogenesis. Consistently, depletion of the DAB component CEP164, as demonstrated
previously 35, 36, or SCLT1, as demonstrated herein, fully abolished DABs and
ciliogenesis (Fig. 7a, b; Supplementary Fig. 15). Specifically, this resulted in
defective initiation (Fig. 7a), since neither CP110 removal nor TTBK2 recruitment
could occur at the mother centriole in SCLT1-/- cells (CRISPR knockout,
Supplementary Fig. 13). However, intriguingly, knockout of the DAM component
FBF1 did not affect ciliogenesis during the same initiation steps. Both CP110 removal
and TTBK2 recruitment occurred normally in FBF1-/- cells (~70%), compared with
WT cells (~80%), and yet cilia were detected in only ~14% of the population (Fig. 7a;
Supplementary Fig. 15). These results suggest that unlike CEP164 or SCLT1, which
are essential for cilia initiation, FBF1 acts downstream of SCLT1 5 and is involved in
cilia maturation and/or maintenance.
The DAM protein FBF1 is associated with ciliary gating of transmembrane
proteins
To explore how FBF1 contributes to cilia maintenance, we examined whether it is
required for ciliary gating. No significant difference in ciliary enrichment of ARL13B
was detected between WT and FBF1-/- cilia (Fig. 7c). However, in striking contrast,
although overexpressed Smoothened was highly concentrated in WT cilia, as reported
previously 37 no such enrichment was observed in FBF1-/- cilia (Fig. 7d, e).
Consistently, the lack of ciliary enrichment was also seen in siFBF1 cells
overexpressing SSTR3, another transmembrane protein known to localize to cilia
(Fig. 7f, g). These results suggest that either the ciliary entry or retention of
transmembrane proteins (Smoothened or SSTR3) is defective in FBF1-/- cilia. We
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repeated the assay in CEP128-/-;C-Nap1-/- cells known to grow fully surfaced cilia 38,
allowing clear distinction between the ciliary membrane and the plasma membrane. In
the presence of FBF1, exogenously expressed Smoothened was highly enriched in
surfaced cilia, but when FBF1 was depleted, as was seen in CEP128-/-;C-Nap1-/-
;siFBF1 cells, the level of Smoothened at surfaced cilia was much lower, although
still detectable (Fig. 7h, i). If this was due to an effect on the entry of transmembrane
proteins, Smo or SSTR3 should have been accumulated close to the DAPs. Therefore,
our findings indicate that, in the absence of FBF1, Smoothened could enter but was
not retained or enriched in the ciliary membrane. A possible reason for the lack of
Smoothened ciliary enrichment could be a defective transition zone, since a previous
study showed that depletion of certain transition zone proteins can inhibit this process
39. However, we excluded this possibility by showing that transition zone proteins
TCTN2, TMEM67, RPGRIP1L, and CEP290 were unaffected in FBF1-/- cells
compared with WT cells, and remained localized at the ciliary base (Fig. 7j). Our
results thus strongly suggest that the DAM component FBF1 performs an essential
role in ciliary gating, particularly of transmembrane proteins (Fig. 7k, l).
Discussion
In summary, using superresolution microscopy, we determined the detailed molecular
architecture of centriolar DAPs and the surrounding structure at the ciliary base,
thereby bridging the gap between structural biology and cell biology, which is
essential for precise functional studies. Our work provides an unprecedented roadmap
to functional investigation based on more than 12 molecular elements at the interfaces
of the centriole and cilium, and the plasma and ciliary membranes. Most importantly,
instead of a pinwheel-like structure based on previous EM studies, our findings reveal
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that DAPs have a conical-shaped architecture composed of nine DABs within an
embedded matrix that we termed the DAM. Both DABs and the DAM have distinct
molecular elements that serve distinct functions. DABs contain CEP164, SCLT1,
CEP89, and CEP83, all of which play an essential role in cilia initiation, whereas the
previously unexplored DAM is filled with IFT molecules and FBF1, which is
intriguingly found to facilitate ciliary gating of transmembrane proteins. As FBF1 is
known to be associated with the apical junction complex that forms the barrier of
epithelia 40, it may also play a similar role at the junction between the ciliary
membrane and the plasma membrane at the base of the ciliary pockets to gate the
ciliary compartment.
Accumulation of IFT88 molecules in the gaps between DABs indicates a unique
molecular arrangement in this region. In order to have protein molecules
concentrated/localized at a specific region of the cell, where they can dynamically
enter/attach and exit/detach, the region must either be gated with a structural barrier,
or alternatively include a scaffold that can bind to these protein molecules with the
required affinity. In either case, dedicated structures must be present in this. In our
current study, we found that FBF1 and IFT88 were concentrated/localized at the space
between DABs, outside of the ciliary compartment, in a region previously thought to
be empty based on EM studies. We thus conclude that, rather than being empty, the
region is equipped with structures that are able to trap protein molecules to form the
DAM. We use the term ‘matrix’ to indicate a surrounding structure or material in
which something develops, rather than pointing to any defined architecture.
This current work provides a functional extension of our previous finding of a
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hierarchical relationship among SCLT1, CEP164, and FBF1 5. While SCLT1 mediates
the recruitment of both FBF1 and CEP164 to centrioles, recruitment of each is
independent of the other. Thus, loss of FBF1 alone is completely different from loss
of SCLT1, which leads to removal of both FBF1 and CEP164 from centrioles. This
previously determined ‘nonlinear’ assembly hierarchy is consistent with the
observations in our current study; Both SCLT1 and CEP164 are components of
DABs, and knockout of either prevents cilia initiation. By contrast, FBF1 is a matrix-
associated protein occupying the space between the blades, and loss of FBF1 affects
cilia maturation/maintenance (the integrity of the ciliary membrane), rather than cilia
initiation.
Our previous genetic studies showed that SCLT1 acts upstream of CEP164, whereas
our superresolution studies showed that SCLT1 is “sandwiched” by the signal from
the ‘blade-like’ CEP164, not immediately obvious how this localization relationship
occurs. The initial recruitment of CEP164, which depends on CEP83 and SCLT1,
may be completely separate from the propagation of CEP164 along the structure that
forms the final localization structure. In particular, CEP164 harbors the coiled-coil
domains known to mediate self-oligomerization. That is, in the absence of
CEP83/SCLT1, CEP164 molecules were not detected at centrioles. However, once
initial CEP164 molecules are recruited, they can recruit other CEP164 molecules
through protein-protein interaction or self-oligomerization processes. A similar
concept can be seen in the PCM organization surrounding the centriole; only the inner
layer of molecules is directly recruited by the centriole, whereas molecules in the
outer layer are recruited through self-assembly or self-oligomerization 41. Our
previous determination of the assembly hierarchy was based on genetics, rather than
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the actual, stepwise, biochemical assembly pathway. The findings of the present study
suggest that the basic framework of the DAP structure can in principle be assembled
in the cytoplasm or in the DAP region initiated by recruited seed molecules to form
the final molecular architecture.
Acknowledgements
We thank Erich Nigg, Christopher Westlake, Steve Caplan and Tang Tang for sharing
reagents. This work was supported by the Ministry of Science and Technology,
Taiwan (Grant No. 103-2112-M-001-039-MY3), Academia Sinica Career
Development Award, Academia Sinica Nano Program, the University of Alabama at
Birmingham (UAB) Hepato/Renal Fibrocystic Diseases Core Center (HRFDCC) Pilot
Award (NIH 5P30DK074038-09) to J.-C. L.; NIH grant GM088253, American
Cancer Society grant RSG-14-153-01-CCG, and the Geoffrey Beene Cancer Research
Center grant to M.-F.B.T.
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9. Kim, S. & Dynlacht, B.D. Assembling a primary cilium. Curr. Opin. Cell Biol. 25, 506‐511 (2013). 10. Anderson, R.G.W. The three‐dimensional structure of the basal body from the rhesus monkey oviduct. J. Cell Biol. 54, 246‐265 (1972). 11. Ye, X., Zeng, H., Ning, G., Reiter, J.F. & Liu, A. C2cd3 is critical for centriolar distal appendage assembly and ciliary vesicle docking in mammals. Proc. Natl. Acad. Sci. U.S.A. 111, 2164‐2169 (2014). 12. Goetz, S C., Liem, K F. & Anderson, K V. The spinocerebellar ataxia‐ associated gene tau tubulin kinase 2 controls the initiation of ciliogenesis. Cell 151, 847‐858 (2012). 13. Nachury, M. et al. A core complex of BBS proteins cooperates with the GTPase Rab8 to promote ciliary membrane biogenesis. Cell 129, 1201 ‐ 1213 (2007). 14. Knödler, A. et al. Coordination of Rab8 and Rab11 in primary ciliogenesis. Proc. Natl. Acad. Sci. U.S.A. 107, 6346‐6351 (2010). 15. Lu, Q. et al. Early steps in primary cilium assembly require EHD1/EHD3‐ dependent ciliary vesicle formation. Nat. Cell Biol. 17, 228‐240 (2015). 16. Caspary, T., Larkins, C.E. & Anderson, K.V. The graded response to Sonic Hedgehog depends on cilia architecture. Dev. Cell 12, 767‐778 (2007). 17. Deane, J.A., Cole, D.G., Seeley, E.S., Diener, D.R. & Rosenbaum, J.L. Localization of intraflagellar transport protein IFT52 identifies basal body transitional fibers as the docking site for IFT particles. Curr. Biol. 11, 1586‐1590 (2001). 18. Spektor, A., Tsang, W.Y., Khoo, D. & Dynlacht, B.D. Cep97 and CP110 suppress a cilia assembly program. Cell 130, 678‐690 (2007). 19. Reiter, J.F., Blacque, O.E. & Leroux, M.R. The base of the cilium: roles for transition fibres and the transition zone in ciliary formation, maintenance and compartmentalization. EMBO Rep. 13, 608‐618 (2012). 20. Chang, B. et al. In‐frame deletion in a novel centrosomal/ciliary protein CEP290/NPHP6 perturbs its interaction with RPGR and results in early‐ onset retinal degeneration in the rd16 mouse. Hum. Mol. Genet. 15, 1847 ‐ 1857 (2006). 21. Wang, W.‐J. et al. CEP162 is an axoneme‐recognition protein promoting ciliary transition zone assembly at the cilia base. Nat. Cell Biol. 15, 591‐ 601 (2013). 22. Rosenbaum, J.L. & Witman, G.B. Intraflagellar transport. Nat. Rev. Mol. Cell Biol. 3, 813‐825 (2002). 23. Rust, M.J., Bates, M. & Zhuang, X.W. Sub‐diffraction‐limit imaging by stochastic optical reconstruction microscopy (STORM). Nat. Methods 3, 793‐795 (2006). 24. Heilemann, M. et al. Subdiffraction‐resolution fluorescence imaging with conventional fluorescent probes. Angew. Chem. Int. Ed. 47, 6172‐6176 (2008). 25. Yang, T.T. et al. Superresolution pattern recognition reveals the architectural map of the ciliary transition zone. Sci. Rep. 5, 14096 (2015). 26. Lau, L., Lee, Yin L., Sahl, Steffen J., Stearns, T. & Moerner, W.E. STED microscopy with optimized labeling density reveals 9‐fold arrangement of a centriole protein. Biophys. J. 102, 2926‐2935 (2012). 27. Lee, Y.L. et al. Cby1 promotes Ahi1 recruitment to a ring‐shaped domain at
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the centriole–cilium interface and facilitates proper cilium formation and function. Mol. Biol. Cell 25, 2919‐2933 (2014). 28. Sillibourne, J.E. et al. Assessing the localization of centrosomal proteins by PALM/STORM nanoscopy. Cytoskeleton 68, 619‐627 (2011). 29. Sonnen, K.F., Schermelleh, L., Leonhardt, H. & Nigg, E.A. 3D‐structured illumination microscopy provides novel insight into architecture of human centrosomes. Biol. Open (2012). 30. Cevik, S. et al. Joubert syndrome Arl13b functions at ciliary membranes and stabilizes protein transport in Caenorhabditis elegans. J. Cell Biol. 188, 953‐969 (2010). 31. Joo, K. et al. CCDC41 is required for ciliary vesicle docking to the mother centriole. Proc. Natl. Acad. Sci. U.S.A. 110, 5987‐5992 (2013). 32. Jensen, V.L. et al. Formation of the transition zone by Mks5/Rpgrip1L establishes a ciliary zone of exclusion (CIZE) that compartmentalises ciliary signalling proteins and controls PIP2 ciliary abundance. EMBO J. 34, 2537‐2556 (2015). 33. Cajanek, L. & Nigg, E.A. Cep164 triggers ciliogenesis by recruiting Tau tubulin kinase 2 to the mother centriole. Proc Natl Acad Sci U S A 111, E2841‐2850 (2014). 34. Wei, Q. et al. Transition fibre protein FBF1 is required for the ciliary entry of assembled intraflagellar transport complexes. 4, 2750 (2013). 35. Graser, S. et al. Cep164, a novel centriole appendage protein required for primary cilium formation. J. Cell Biol. 179, 321‐330 (2007). 36. Schmidt, K.N. et al. Cep164 mediates vesicular docking to the mother centriole during early steps of ciliogenesis. J. Cell Biol. 199, 1083‐1101 (2012). 37. Corbit, K.C., Aanstad, P., Singla, V. & Norman, A.R. Vertebrate Smoothened functions at the primary cilium. Nature 437, 1018 (2005). 38. Mazo, G., Soplop, N., Wang, W.‐J., Uryu, K. & Tsou, M.‐F.B. Spatial control of primary ciliogenesis by subdistal appendages alters sensation‐associated properties of cilia. Dev. Cell 39, 424‐437 (2016). 39. Garcia‐Gonzalo, F. et al. A transition zone complex regulates mammalian ciliogenesis and ciliary membrane composition. Nat Genet 43, 776 ‐ 784 (2011). 40. Sugimoto, M. et al. The keratin‐binding protein Albatross regulates polarization of epithelial cells. J. Cell Biol. 183, 19‐28 (2008). 41. Mennella, V. et al. Subdiffraction‐resolution fluorescence microscopy reveals a domain of the centrosome critical for pericentriolar material organization. Nat Cell Biol 14, 1159‐1168 (2012). 42. Berbari, N., Johnson, A., Lewis, J., Askwith, C. & Mykytyn, K. Identification of ciliary localization sequences within the third intracellular loop of G protein‐coupled receptors. Mol Biol Cell 19, 1540 ‐ 1547 (2008). 43. Mali, P. et al. RNA‐guided human genome engineering via Cas9. Science 339, 823‐826 (2013).
17 bioRxiv preprint doi: https://doi.org/10.1101/193474; this version posted September 25, 2017. The copyright holder for this preprint (which was not certified by peer review) is the author/funder. All rights reserved. No reuse allowed without permission.
Figure Legends
Figure 1 Hierarchical and patterned radial localization of proteins at the distal
appendage (DAP) region revealed by superresolution microscopy.
(a) Definitions of view angles (axial and lateral) and orientations of molecular
distributions (radial, circumferential, and longitudinal) on a mother centriole. (b)
dSTORM superresolution images showing the distribution patterns of various DAP,
subdistal appendage, and centriolar proteins in order of radial size, which are not
resolvable by widefield (WF) imaging. (c) Axial view EM image overlaid with single-
color superresolution images of CEP164, SCLT1, CEP83, Cby1, CP110, and C2CD3,
illustrating that CP110 overlaps with axonemal microtubules, whereas SCLT1 and
CEP164 are localized near the tips of highly electron dense branches of the EM image
that represent DAPs. (d) Mean diameter analysis revealing dimensional differences
among ciliary proteins in (b), where the diameters of CP110, CEP97, CEP162, and
CEP290 are similar to that of the axoneme measured by EM (dashed line). CEP164
(o) and CEP164 (i) represent the outer and inner diameters of CEP164, respectively.
Bars = 200 nm (b) and 100 nm (c).
Figure 2 Angular distributions of CEP83, CEP89, SCLT1, FBF1, and CEP164
revealed by two-color axial view dSTORM imaging.
(a-e) Two-color axial view images of (a) CEP83-SCLT1, (b) SCEP89-CEP164, (c)
CEP164-SCLT1, (d) CEP164-FBF1, and (e) FBF1-SCLT1 pairs. CEP83 and SCLT1
(a) and CEP89 and CEP164 (b) align well with each other. SCLT1 likely resides
within each ‘blade-like’ distribution of CEP164 signals (c), whereas the angular
positions of FBF1 puncta are different from those of CEP164 puncta (d) and SCLT1
puncta (e). White bars mark SCLT1 in (a) or CEP164 in (be); circles mark SCLT1 in
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(c) or FBF1 in (de); triangles mark SCLT1 in (d). (f) Angular analysis in the
circumferential direction shows that CEP83, CEP89, SCLT1, and CEP164 exhibit
similar angular distributions, while FBF1 is arranged alternately with SCLT1 or
CEP164, suggesting it fills the gaps between adjacent CEP164 puncta. (g) Model of
the localization of DAP proteins. FBF1 occupies the gaps between DAP blades,
whereas C2CD3 localizes to the centriolar lumen. Bars (ae) = 200 nm.
Figure 3 3D spatial map of distal appendage (DAP)-associated proteins revealing
unique FBF1 localization at the gaps between DAP blades.
(a) Three representative two-color dSTORM images from a lateral view of each pair
of DAP proteins illustrates their relative longitudinal positions. Pink and green
arrowheads depict the heights of FBF1 and SCLT1, respectively. (b) Representative
images of DAP proteins are aligned and combined according to their average
longitudinal positions. The magnified image (inset) demonstrates the mapping of DAP
proteins on the tilted arrangement of the DAP structure (dashed line). (c)
Superresolved images of the subdistal appendage protein ODF2 reveal its two-layered
localization at a far lower location than that of SCLT1. (d) Merged superresolution
image in (b) overlaid with a longitudinally sliced EM micrograph. CEP83, SCLT1,
and FBF1 are well aligned with the electron density of the slanted DAPs. (e) CP110
and CEP97 are localized longitudinally, similar to SCLT1, but with a narrower lateral
width. (f) Combined relative localization of DAP-associated proteins in radial and
longitudinal directions, revealing the envelope of the slanted arrangement of a DAP
blade (dotted line). (g) A 3D computational model illustrating the positioning of
proteins at the DAP region with a backbone structure based on EM 10. (h) CEP164,
CEP83, SCLT1, and the distal layer of CEP89 mapped onto a blade of the 3D-
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reconstructed DAPs. (i) FBF1 localizes to gaps between two adjacent DAP blades,
possibly at the proximal boundary of the ciliary pocket. Bars a, b (left), c, and e = 200
nm; b (inset) = 50 nm.
Figure 4 Localization of other DAP-related proteins ARL13B, EHD1, and TTBK2.
(a) During the early stages of cilia growth, ARL13B is located adjacent to FBF1
(arrows), as confirmed by correlation analysis in (b). (c) ARL13B becomes localized
to cilia during or after ciliogenesis with an exclusion zone approximately at the
proximal transition zone. (d) The vesicle-forming protein EHD1 is localized to a
peripheral region wider than that of ARL13B, presumably on the membrane of the
ciliary pocket. Axial (e, f) and lateral (g) views of the distribution of TTBK2 with and
without serum, showing a broader and more enriched TTBK2 distribution in both
radial and longitudinal directions after serum starvation. Bars = 200 nm.
Figure 5 Localization of IFT88 revealing the existence of a distal appendage matrix
(DAM) between distal appendage blades (DABs).
(a) Three representative axial view dSTORM images of IFT88 reveal a broad angular
distribution without a clear 9-fold symmetric pattern. (b) Two representative sets of
two-color images showing IFT88 spanning a longitudinal range relative to the
location of SCLT1 (marked with a dashed line). The solid line in (c) depicts the
distribution in the lower panel in (b), while the dotted line illustrates normalized
distributions averaged over nine data pairs. (d, e) 3D STORM imaging of IFT88
showing the tapered distribution of IFT88 that is wider at the distal end of the DAP
region. (f) 3D two-color superresolution image revealing IFT88 proteins in the gaps
between neighboring SCLT1 regions, as shown in the lower image (derived from the
20 bioRxiv preprint doi: https://doi.org/10.1101/193474; this version posted September 25, 2017. The copyright holder for this preprint (which was not certified by peer review) is the author/funder. All rights reserved. No reuse allowed without permission.
upper-right image) in which nine SCLT1 puncta are numbered accordingly. (g) The
region between adjacent DABs is designated as the DAM. (h) Model showing the
localization of FBF1 and IFT88 at the DAM. (i) Two-color imaging of IFT88-FBF1
shows the angular coincidence between IFT88 and FBF1 at the DAM. (j) Histogram
of angular spacing between IFT88 and FBF1. (k, l) The radial distribution of IFT88
was reduced in FBF1-/- cells. (m) Model showing a functional role of FBF1 in
maintaining the DAM spatial arrangement. Bars = 200 nm for all, except f (100 nm).
Figure 6 DAB components mediating the structural arrangement of the DAM.
(a, b) Axial view dSTORM images reveal the distribution of (a) CEP164 and (b)
FBF1 at the mother centrioles upon CRISPR/Cas9 FBF1 knockout and CEP164
siRNA silencing, respectively. Note that the FBF1 ring formed upon CEP164
knockdown becomes smaller than that of wild-type (WT) cells (b, right panel).
(c, d) Statistical analysis of the radial distribution of CEP164 and FBF1 upon
perturbation. The size of CEP164 in FBF1-/- cells is comparable to that in WT cells
(c), whereas the size of FBF1 is reduced upon CEP164 knockdown (d). (e) Model
showing that the arrangement of FBF1 is no longer maintained at the outer region of
the DAM in the absence of CEP164. (f) 3D model based on all axial and lateral
superresolution images illustrating the molecular architecture on the framework of
DABs, DAM, axoneme, and ciliary membrane. (g) Close-up view of the DAP region
suggesting that FBF1 potentially interfaces the ciliary membrane and the DAM.
Figure 7 The DAB protein SCLT1 in ciliogenesis vs. the DAM protein FBF1 in
transmembrane protein gating.
(a) Deletion of SCLT1 fully inhibits ciliogenesis during the initiation stage, since
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neither CP110 removal nor TTBK2 recruitment occurred at the mother centriole.
However, both CP110 removal and TTBK2 recruitment were observed in FBF1-/-
cells. White boxes highlight the areas at the basal bodies or centrioles shown in the
insets. (b) EM micrographs showing the absence of DABs in SCLT1-/- cells and
visible DABs in wild-type cells (arrows). Subdistal appendages (double arrows)
remain present in SCLT1-/- cells. (c) ARL13B is enriched in both WT and FBF1-/-
cilia. (d, e) Overexpressed Smoothened was highly concentrated in WT cilia but was
not enriched in FBF1-/- cilia. The insets in (d) show enlarged images of the boxed
region. (e) Smo-GFP intensities relative to cytoplasmic levels. (f, g) SSTR3-GFP
signals at cilia were lower in siFBF1 cells. (h, i) Overexpressed Smoothened is
abundant in surfaced cilia in CEP128-/-;C-Nap1-/- cells, but less enriched in surfaced
cilia in CEP128-/-;C-Nap1-/-;siFBF1 cells, suggesting Smoothened was not retained in
FBF1-depleted cilia. (j) Enrichment of transition zone proteins TCTN2, TMEM67,
RPGRIP1L, and CEP290 was unaffected in FBF1-/- cells compared with WT cells. (k,
l) Model showing that FBF1 plays a role in gating transmembrane proteins,
presumably at the junction between the ciliary membrane and the plasma membrane.
Bar in (b) = 100 nm.
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Figure 1
Figure 2
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Figure 3
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Figure 4
Figure 5
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Figure 6
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Figure 7
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METHODS
Antibodies
Detailed information about the primary antibodies against CEP164, FBF1, SCLT1,
CEP89, CEP83, Cby1, CP110, CEP97, CEP162, CEP290, C2CD3, IFT88, TTBK2,
ODF2, EHD1, ARL13B, and Centrin is provided in Supplementary Table 2.
Antibodies conjugated to Alexa Fluor 488 (mouse A21202 and rabbit A21206;
Thermo Fisher Scientific, Waltham, MA, USA) and Alexa Fluor 647 (anti-mouse
A21236, anti-rabbit A21245, anti-rat A21247, and anti-goat A21447; Thermo Fisher
Scientific) were used as secondary antibodies. Cy3B-conjugated secondary antibody
was custom-made by conjugating Cy3B maleimide (PA63131, GE, Pittsburgh, PA,
USA) to IgG antibodies (rabbit 711-005-152 and rat 712-005-153; Jackson
ImmunoResearch, West Grove, PA, USA).
Cell culture and transfection
Human retinal pigment epithelial cells (hTERT RPE-1) were seeded on poly-L-lysine-
coated coverslips and cultured at 37C under 5% CO2 in DMEM/F-12 mixture
medium (1:1; 11330-032, Gibco, Thermo Fisher Scientific) supplemented with 10%
fetal bovine serum (FBS) and 1% penicillin-streptomycin. HEK293T cells were
cultured in DMEM with 10% FBS and 1% penicillin-streptomycin. Cilium formation
in RPE-1 cells was induced by serum starvation for 2448 h. Lentiviral stocks were
prepared according to standard procedures (PT-5144-1, Clontech, Mountain View,
CA, USA). RNAi-mediated knockdown of human FBF1 in RPE-1 cells was
performed using 12.5 μM of three different siRNA duplex oligonucleotides (Thermo
Fisher, ambion silencer select) targeting the sequences 5′-
GGGAGAACCAGGCTCCAAATT-3′ and 5′-GCATGGATGCTGATATCTTTT-3′,
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and 5′-TGAGAAGCCTGTTAAACTATT-3′. siRNA oligos were delivered using
Lipofectamine RNAiMAX Reagent (13778150, Thermo Fisher). Human CEP89
(NM_032816) depletion was carried out using a combination of five different
lentiviral shRNAs (RNAi core, Academia Sinica, Taiwan) with the following
targeting sequences: sh1, 5’-GCACGTCAAAGATATACAGAA-3’; sh 2, 5’-
CAGGAGTCATTTCAAACACAT-3’; sh 3, 5’-GAATGTAGAACTCAGTCGATA-
3’; sh 4, 5’-CCAGATATAACTGGTAGAGCA-3’, and sh 5, 5’-
GTAACGTTTCCGATTGTGAAA-3’. All lentiviruses were generated by transient
cotransfection of HEK293T cells with packaging and envelope vectors (PT-5144-1,
Clontech, Mountain View, CA, USA) using a TransIT-293 Transfection Reagent Kit
(MIR 2700, Mirus, Madison, WI, USA). Stable expression of human CEP89 in RPE1
cells was performed through lentiviral transduction and selection using 5 mg/mL
puromycin. To induce ciliary growth, RPE-1 was starved of serum for 48 h. SSTR3-
GFP (#35623, Addgene 42) was cloned into the pLVX-Tight-Puro vector (Clontech)
for tet-inducible expression, as was Smoothened-GFP (described previously 38). For
Smo-GFP/SSRT3-GFP, lentivirus production and infection were performed as
described previously 5 using psPax2 (packaging, #12260, Addgene), pCMV-VSV-G
(envelope, #8454, Addgene), and pLVXTet-On Advance (Clontech) plasmids. Cells
were infected simultaneously with lentiviral vectors for rttA expression (Tet-On
advance) and Smo-GFP (or SSTR3-GFP). Clonal populations displaying uniform
transgene expression were isolated prior to siRNA experiments. Tet induction was
performed with 1 g/mL doxycycline for 48 h (during serum starvation).
CRISPR construction and generation of SCLT1-/- and FBF1-/- cells
RNA-guided targeting of genes in human cells was achieved through coexpression of
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the Cas9 protein with gRNAs using reagents prepared by the Church research group
43, which are available from Addgene (http://www.addgene.org/crispr/church/).
Targeting sequences for SCLT1 and FBF1 were 5’-GGTGTCTGCCCATGGAAGAG-
3’ and 5’-GGCATGGATGCTGATATCTT-3’, respectively, and these were cloned into
the gRNA cloning vector (#41824, Addgene, Cambridge, MA, USA) via the Gibson
assembly method (New England Biolabs, Ipswich, MA, USA) as described previously
43. Pure SCLT1 and FBF1 knockout cell lines were then established through clonal
propagation from single cells. For genotyping, the following PCR primers were used:
5’-ACGCGGATCCAACGTAAGTGTTAGAATGCT-3’ and 5’-
CTAGTCTAGAAGCTAGGTGCATTTAACAAT-3’ for SCLT1 alleles, and 5’-
ACGCGGATCCCCCGAGACCAGCCACCTAGG-3’ and 5’-
CTAGTCTAGACCTTTCTCACTGGCTACTGA-3’ for FBF1 alleles. PCR products
were cloned and sequenced.
Immunofluorescence
Cells on glass coverslips were fixed with either 4% paraformaldehyde at room
temperature or methanol at -20C after one wash with PBS. Cells were then
permeabilized with 0.1% PBST (PBS with 0.1% Triton X-100) for 10 min and
blocked with 3% BSA for 30 min. Primary antibodies were diluted to optimized
concentrations with 0.1% BSA in PBST and applied to cells. After a 1 h incubation
with antibody, cells were washed three times with 0.1% PBST to remove unbound
antibodies, then incubated with optimally diluted secondary antibodies for 1 h. Cells
were finally washed three times with 0.1% PBST and stored in PBS with sodium
azide at 4C until later use.
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dSTORM imaging and analysis
Superresolution imaging was performed with a modified inverted microscope (Eclipse
Ti-E, Nikon, Tokyo, Japan). Three light sources were merged onto a laser merge
module (ILE, Spectral Applied Research, Richmond Hill, Ontario, Canada), which
was integrated with individual controllers. Beams from a 637 nm laser (OBIS 637 LX
140 mW, Coherent, Santa Clara, CA, USA), a 561 nm laser (Jive 561 150 mW,
Cobolt, Solna, Sweden) and a 405 nm laser (OBIS 405 LX 100 mW, Coherent) were
homogenized (Borealis Conditioning Unit, Spectral Applied Research) and focused
on the back aperture of a 100 1.49 NA oil immersion objective (CFI Apo TIRF,
Nikon) for wide-field illumination of samples. A perfect focusing system (PFS,
Nikon) was used to minimize z-axis drift. The 637 nm and 561 nm laser lines were
operated at a high intensity of ~15 kW/cm2 to quench most of the fluorescence from
Alexa 647. A weak 405 nm beam was used to activate a portion of the dyes that were
converted from a ground state to an excited state. The collected fluorescence was
cleaned by a single-band emission filter and imaged on an EMCCD camera (Evolve
512 Delta, Photometrics, Tucson, AZ, USA) with an overall pixel size of 93 nm.
Fiducial markers (Tetraspeck, Thermo Fisher) were used to correct the drift. Typically,
10,00020,000 frames (for 2D dSTORM) and ~50,000 frames (for 3D dSTORM)
were recorded at a rate of 5067 fps. Individual single-molecule peaks were then real-
time localized using a MetaMorph Superresolution Module (Molecular Devices,
Sunnyvale, CA, USA), based on a wavelet segmentation algorithm. Superresolution
images in figures were cleaned with a Gaussian filter with a radius of 0.61 pixels.
For single-color imaging of Alexa 647, signals were filtered with a bandpass filter
(700/75, Chroma, Bellows Falls, VT, USA); for two-color imaging, the Alexa 647
channel was first recorded, then the Cy3B channel was acquired with the
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corresponding filter (593/40, Chroma). For 3D imaging, a cylindrical lens with a focal
length of 25 cm was added prior to the camera to create astigmatism, from which the
z position was determined through calibration. The point spread function (PSF)-based
calibration curve was generated by scanning a fluorescent bead in the z direction and
calculating the elliptical widths (wx and wy).
dSTORM imaging was performed in an imaging buffer containing TN buffer at pH
8.0, and an oxygen-scavenging system consisting of 60100 mM mercaptoethylamine
(MEA) at pH 8.0, 0.5 mg/mL glucose oxidase, 40 g/mL catalase, and 10% (w/v)
glucose (Sigma-Aldrich, St. Louis, MO, USA). Lateral position drift was measured
during the acquisition and corrected by frame-by-frame correlation of the fiducial
markers with ImageJ. Chromatic aberration between long and short wavelength
channels was compensated with a customized algorithm relocating each pixel of a 561
nm image to its targeted position in the 647 nm channel with a predefined correction
function obtained by parabolic mapping of multiple calibration beads. For axial
imaging of DAPs, a ring pattern of labeled SCLT1 or FBF1 was usually sought to
identify those with appropriate orientations. For lateral imaging, a rod-like pattern of
SCLT1 or FBF1 was used to assure a lateral view of the centriole-cilium.
To determine the diameter of DAPs and their associated proteins shown in Fig. 1b, the
position of individual superresolved puncta forming a ring-shaped pattern was first
measured and then fitted with a circle to estimate its diameter. For CEP164, Cby1,
IFT88, and TTBK2, where their distributions were relatively dispersed, their radial
positions were described with a radius defined as the distance between the puncta and
the center, yielding a radius histogram that was fitted with a Gaussian or mixed
32 bioRxiv preprint doi: https://doi.org/10.1101/193474; this version posted September 25, 2017. The copyright holder for this preprint (which was not certified by peer review) is the author/funder. All rights reserved. No reuse allowed without permission.
Gaussian curve (Supplementary Fig. 1). To determine how DAP proteins aligned with
each other (Fig. 2ae), angular analysis was performed. Briefly, the angular position
of each DAP punctum was recorded, and angle differences between each pair of
puncta for two protein species indicated how they were arranged relative to each
other. The concept is described in detailed in Supplementary Fig. 4. To align
dSTORM and EM images in Fig. 3d, FBF1 signals of dSTORM images were placed
at the tip of the high electron density signals of slanted DAPs in EM images.
Transmission electron microscopy
RPE-1 cells grown on coverslips made of Aclar film (Electron Microscopy Sciences,
Hatfield, PA, USA) were fixed in 4% paraformaldehyde and 2.5% glutaraldehyde
with 0.1% tannic acid in 0.1 M sodium cacodylate buffer at room temperature for 30
min, postfixed in 1% OsO4 in sodium cacodylate buffer for 30 min on ice, dehydrated
in a graded series of ethanol, infiltrated with EPON812 resin (Electron Microscopy
Sciences), and embedded in the resin. Serial sections (~90 nm thickness) were cut on
a microtome (Ultracut UC6; Leica, Wetzlar, Germany) and stained with 1% uranyl
acetate and 1% lead citrate. Samples were examined on a transmission electron
microscope (Tecnai G2Spirit Twin, FEI, Hillsboro, OR, USA).
3D printing model
To reconstruct a 3D model of DAPs, we first drew the DAP structure using 3D CAD
design software (SOLIDWORKS, Waltham, MA, USA) then fabricated a solid model
with a 3D printer (Form 1+, Formlabs, Somerville, MA, USA). The dimensions of the
model were based on the localization positions obtained from the superresolution
results in Fig. 1 (A, C, D, E), Fig. 2 (A, E, F), Fig. 3 (A to C, I, N, O, P, Q), and on
33 bioRxiv preprint doi: https://doi.org/10.1101/193474; this version posted September 25, 2017. The copyright holder for this preprint (which was not certified by peer review) is the author/funder. All rights reserved. No reuse allowed without permission.
information from previous studies 7, 10. For the 3D printed model, DAP proteins in
each ‘blade’ were printed as spherical shapes with their centers at the mean positions,
then painted in the appropriate colors. The printed model, including microtubule
triplets and doublets, DABs, transition zone, and ciliary pocket, has an overall size of
85 mm in width (diameter) by 155 mm in height at a 150,000:1 scale. To properly
place FBF1 in the gaps between two adjacent DAPs, an auxiliary annular scaffold,
which does not exist in the actual complex, was added.
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